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Abstract:

Magnetic particle imaging (MPI) is an emerging imaging modality that enables the direct mapping of iron oxide nanoparticle tracers.  In MPI, the development of tailored magnetic nanoparticle tracers is paramount to achieving high sensitivity and good spatial resolution. This talk will provide a general overview of the progress in MPI tracer development over the past decade, and will also focus on emerging directions and new opportunities for iron oxide-based tracer design and applications. The presentation will cover magnetic nanoparticle relaxation in MPI and discuss key aspects to consider in tailoring tracers for MPI applications. Emphasis will be given on how structural changes (size, composition, shape, surface chemistry) and inter-particle interactions affect the MPI signal generation process. Moreover, the presentation will discuss emerging research directions in color-MPI (cMPI) and MPI-guided hyperthermia (hMPI).

Abstract

Lignin remains one of the most significant barriers to the efficient utilization of lignocellulosic substrates, in processes ranging from ruminant digestibility to indus­trial pulping, and in the current focus on biofuels production. Inspired largely by the recalcitrance of lignin to biomass processing, plant en­gineering has routinely sought to alter lignin quantity, composition, and structure by exploiting the inherent plasticity of lignin biosynthesis.



More recently, researchers are attempting to strategically design plants for increased degradability by incorporating monomers that lead to a lower degree of polymerization, reduced hydrophobicity, fewer bonds to other cell wall constituents, or novel chemically labile linkages in the polymer backbone.1,2 The incorporation of value-added structures could also help valorize lignin. Designer lignins may satisfy the biological requirement for lignification in plants while improving the overall efficiency of biomass utilization. Although possibilities abound, maintaining plant health is paramount and, ultimately, the plants themselves will dictate which of these approaches can be tolerated.



One such method, via the so-called ‘zip-lignin’ approach, is showing particular promise.3-5 Poplar has been engineered to incorporate monolignol ferulate conjugates into the lignification process, by using an exotic transferase gene, FMT, and a xylem-specific promoter.5 This results in the introduction of readily cleavable ester linkages into the backbone of the polymer, and delivers significantly improved processing. Various applications for which these altered trees appear superior are only just beginning to be explored. Now that we have sensitive methods for determining if/when/whether plants are making monolignol ferulate conjugates and using them for lignification, it appears that Nature herself may have already been exploring this avenue.6 In attempting to modify monocot lignins, another transferase appears to be useful for improving cell wall digestibility. This involves p-coumaroylation of the lignin. Although this occurs naturally in monocots, it is not evident in dicots, but providing dicots with that pathway has interesting implications.7,8

Nature continues to inspire us with new avenues toward lignin modification that have potential value for various processes. For example, the ramifications of finding that grasses are using tricin, a flavone from beyond the monolignol biosynthetic pathway, to start lignin chains are interesting; tricin itself is valuable, and plants with tricin knocked out have higher lignin and lower CW digestibility.9-13 

1.    Mottiar Y, Vanholme R, Boerjan W, Ralph J, & Mansfield SD (2016) Designer lignins: harnessing the plasticity of lignification. Current Opinion in Biotechnology 37(1):190-200.



2.    Rinaldi R, Jastrzebshi R, Clough MT, Ralph J, Kennema M, Bruijnincx PCA, & Weckhuysen BM (2016) Paving the way for lignin valorisation: Recent advances in bioengineering, biorefining and catalysis. Angewandte Chemie (International Edition) 55(29):8164-8215.



3.    Grabber JH, Hatfield RD, Lu F, & Ralph J (2008) Coniferyl ferulate incorporation into lignin enhances the alkaline delignification and enzymatic degradation of maize cell walls. Biomacromolecules 9(9):2510-2516.



4.    Ralph J (2010) Hydroxycinnamates in lignification. Phytochemistry Reviews 9(1):65-83.



5.    Wilkerson CG, Mansfield SD, Lu F, Withers S, Park J-Y, Karlen SD, Gonzales-Vigil E, Padmakshan D, Unda F, Rencoret J, & Ralph J (2014) Monolignol ferulate transferase introduces chemically labile linkages into the lignin backbone. Science 344(6179):90-93.



6.    Karlen SD, Zhang C, Peck ML, Smith RA, Padmakshan D, Helmich KE, Free HCA, Lee S, Smith BG, Lu F, Sedbrook JC, Sibout R, Grabber JH, Runge TM, Mysore KS, Harris PJ, Bartley LE, & Ralph J (2016) Monolignol ferulate conjugates are naturally incorporated into plant lignins. Science Advances 2(10):e1600393.



7.    Smith RA, Gonzales-Vigil E, Karlen SD, Park J-Y, Lu F, Wilkerson CG, Samuels L, Mansfield SD, & Ralph J (2015) Engineering monolignol p-coumarate conjugates into Poplar and Arabidopsis lignins. Plant Physiology 169(4):2992-3001.



8.    Sibout R, Le Bris P, Legee F, Cezard L, Renault H, & Lapierre C (2016) Structural redesigning Arabidopsis lignins into alkali-soluble lignins through the expression of p-coumaroyl-CoA:monolignol transferase PMT. Plant Physiology 170(3):1358-1366.



9.    del Río JC, Rencoret J, Prinsen P, Martínez ÁT, Ralph J, & Gutiérrez A (2012) Structural characterization of wheat straw lignin as revealed by analytical pyrolysis, 2D-NMR, and reductive cleavage methods. Journal of Agricultural and Food ����vlog�ٷ���� 60(23):5922-5935.



10. Lan W, Lu F, Regner M, Zhu Y, Rencoret J, Ralph SA, Zakai UI, Morreel K, Boerjan W, & Ralph J (2015) Tricin, a flavonoid monomer in monocot lignification. Plant Physiology 167(4):1284-1295.



11. Lan W, Rencoret J, Lu F, Karlen SD, Smith BG, Harris PJ, del Rio JC, & Ralph J (2016) Tricin-lignins: Occurrence and quantitation of tricin in relation to phylogeny. The Plant Journal 88(6):1046-1057.



12. Lan W, Morreel K, Lu F, Rencoret J, del Rio JC, Voorend W, Vermerris W, Boerjan W, & Ralph J (2016) Maize tricin-oligolignol metabolites and their implications for monocot lignification. Plant Physiology 171(2):810-820.



13. Eloy NB, Voorend W, Lan W, Cesarino I, Vanholme R, de Lyra Soriano Saleme M, Goeminne G, Pallidis A, Morreel K, Nicomedes J, Ralph J, & Boerjan W (2017) Silencing chalcone synthase in maize impedes the incorporation of tricin into lignins and increases lignin content. Plant Physiology 173:998-1016.

Molecular doping is now increasingly used to control the electronic and electrical properties of

organic semiconductors, lower contact resistance, enhance bulk conductivity and carrier mobility,

and create higher performance devices. In this talk, I review processes and options for bulk and

interface doping in molecular and polymer semiconductors, and the roles electron spectroscopy and

carrier transport measurements play in defining key issues. Recent results on p- and n-doping are

described. P-doping using 2,2′-(perfluoronaphthalene-2,6-diylidene)dimalononitrile (F6-TCNNQ)

is tested on two relatively challenging hole transport materials, 2,2′,7,7′-Tetrakis(N,Ndiphenylamino)-

9,9-spirobifluorene (Spiro-TAD) and tris(4-carbazoyl-9-ylphenyl)amine (TCTA)

[1]. The full electronic parameters of these molecules are determined via combinations of electron

spectroscopies. Temperature-dependent transport measurements are done to establish film

conductivity and hole hopping activation energy as a function of dopant concentration. We then

turn to the challenge of n-doping very low electron affinity (EA) electron transport layers (ETL), an

issue critical to OLEDs. We look at the air-stable dimer of (pentamethylcyclopentadienyl)(1,3,5-

trimethylbenzene)ruthenium ([RuCp*Mes]2) [2], and use it to n-dope phenyldi(pyren-2-

yl)phosphine oxide (POPy2) (EA = 2.1 eV). We demonstrate that photo-activation of the cleavable

dimeric dopant results in kinetically stable and efficient n-doping of the host semiconductor, whose

reduction potential is beyond the thermodynamic reach of the dimer’s effective reducing strength

[3]. We demonstrate the use of this doped ETL to fabricate high-efficiency organic light-emitting

diodes.

[1] F. Zhang and A. Kahn, Adv. Funct. Mat. 28, 1703780 (2018)

[2] G. Song, S.-B. Kim, S. Mohapatra, Y. Qi, T. Sajoto, A. Kahn, S. R. Marder and S.

Barlow, Adv. Mat. 24, 699 (2012)

[3] X. Lin, B. Wegner, K. M. Lee, M. Fusella, F. Zhang, K. Moudgil, S. Barlow, S. R.

Marder, N. Koch and A. Kahn, Nature Materials, 16, 1209 (2017)

Iron enzymes activate dioxygen for a large number of biomedically, agriculturally, and environmentally important oxidation reactions. Among those that use non-heme mono-iron cofactors, protein ligand sets as minimal as a pair of cis histidines enable the cofactor to coordinate and approximate as many as three different substrates, leading to an astounding array of different reaction types. Many of these reactions install key functional groups in lucrative natural-product drugs. Over the last 15 years, we have established the intermediacy of iron(IV)-oxo (ferryl) complexes in the reactions of several such enzymes with a range of distinct outcomes.¹ In general, the ferryl intermediates generate substrate radicals by abstracting hydrogen (H•) from unactivated aliphatic carbons,^2-6 initiating formation of new C–O,^2-4 ��–C��/����,^5,6 ��–S,⁷ ��–N,⁸ or C–C bonds.⁹ Hydroxylation is the default outcome, as it results from coupling between the carbon radical and the necessarily adjacent Fe(III)-coordinated oxygen that just generated it (termed “oxygen rebound” by Groves). Thus, the first imperative for enzymes that mediate outcomes other than hydroxylation is to avoid the default rebound, which generally has a low activation barrier. In our current work, we are defining the manifold of other pathways by which the substrate radical can decay and the strategies by which a given protein scaffold (or, in rare cases, the substrate itself) specifies a reaction channel. A generally important parameter is the disposition of the substrate relative to the cofactor,¹⁰ which appears to be controlled not only by the substrate pocket but also by the geometry of the ferryl complex (oxo trans to either one of the two conserved histidine ligands).¹¹ It has therefore been important to have tools to visualize intermediates (especially the ferryl complexes) through the reaction sequences.^12,13 I will summarize several of the Penn State team’s recent successes in defining reaction pathways and explaining control of outcome in this versatile enzyme family.  

1.  Krebs, C., Galonic, D.; Walsh, C. T.; Bollinger, J. M., Jr. "Non-Heme Fe(IV)-Oxo Intermediates," Acc. Chem. Res., 2007, 40, 484-492.

2.  Price, J. C.; Barr, E. W.; Tirupati, B.; Bollinger, J. M., Jr.; Krebs, C.; “The First Direct Characterization of a High-Valent Iron Intermediate in the Reaction of an aKetoglutarate-Dependent Dioxygenase: A High-Spin Fe(IV) Complex in Taurine:a-Ketoglutarate Dioxygenase (TauD) from Escherichia coli,” Biochemistry, 2003, 42, 7497-7508.

3.  Price, J. C.; Barr, E. W.; Glass, T. E.; Krebs, C.; Bollinger, J. M., Jr.; “Evidence for Hydrogen Abstraction from C1 of Taurine by the High-Spin Fe(IV) Intermediate Detected during Oxygen Activation by Taurine:a-Ketoglutarate Dioxygenase (TauD),” J. Am. Chem. Soc., 2003, 125, 13008-13009.

4.  Hoffart, L. M.; Barr, E. W.; Guyer, R. B.; Bollinger, J. M., Jr.; Krebs, C. “Direct spectroscopic detection of a C-H-cleaving high-spin Fe(IV) complex in a prolyl-4-hydroxylase,” Proc. Natl. Acad. Sci. USA, 2006, 103, 14738-14743.

5.  Galonic, D. P.; Barr, E. W.; Walsh, C. T.; Bollinger, J. M., Jr.; Krebs, C. “Two Interconverting Fe(IV) Intermediates in Aliphatic Chlorination by the Halogenase CytC3,” Nat. Chem. Biol., 2007, 3, 113-116.

6.  Matthews, M. L.; Krest, C. M.; Barr, E. W.; Vaillancourt, F. H.; Walsh, C. T.; Green, M. T.; Krebs, C.; Bollinger, J. M., Jr. “Substrate-Triggered Formation and Remarkable Stability of the C-H Bond-Cleaving Chloroferryl Intermediate in the Aliphatic Halogenase, SyrB2,” Biochemistry, 2009, 48, 4331-4343.

7.  Tamanaha, E. Y.; Zhang, B.; Guo, Y.; Chang, W.-c.; Barr, E. W.; Xing, G.; St. Clair, J.; Ye, S.; Neese, F.; Bollinger, J. M., Jr.; Krebs, C. “Spectroscopic evidence for the two C-H-cleaving intermediates of Aspergillus nidulans isopenicillin N synthase," J. Am. Chem. Soc. 2016, 138, 8862-8874.

8.  Matthews, M.L.; Chang, W.-c.; Layne, A.P.; Miles, L.A.; Krebs, C.; Bollinger, J. M., Jr. “Direct Nitration and Azidation of Aliphatic Carbons by an Iron-dependent Halogenase,” Nat. Chem. Biol. 2014, 10, 209-215.

9.  Dunham, N. P.; Chang, W.-c.; Mitchell, A. J.; Martinie, R. J.; Zhang, B.; Bergman, J. A.; Rajakovich, L. J.; Wang, B.; Silakov, A.; Krebs, C.; Boal, A. K.; Bollinger, J. M., Jr. (2018) "Two Distinct Mechanisms for C–C Desaturation by Iron(II)- and 2-(Oxo)glutarate-Dependent Oxygenases: Importance of α-Heteroatom Assistance," J. Am. Chem. Soc., in review.

10. Matthews, M. L.; Neumann, C. S.; Miles, L. A.; Grove, T. L.; Booker, S. J.; Krebs, C; Walsh, C. T.; Bollinger, J. M., Jr. "Substrate positioning controls the partition between halogenation and hydroxylation in the aliphatic halogenase, SyrB2," Proc. Natl. Acad. Sci. USA, 2009, 106, 17723-17728.

11. Martinie, R. J.; Livada, J.; Chang, W-c.; Green, M. T.; Krebs, C.; Bollinger, J. M., Jr.; Silakov, A. "Experimental Correlation of Substrate Position with Reaction Outcome in the Aliphatic Halogenase, SyrB2," J. Am. Chem. Soc. 2015, 137, 6912-6919.

12. Martinie, R. J.; Pollock, C. J.; Matthews, M. L.; Bollinger, J. M., Jr.; Krebs, C; Silakov, A. “Vanadyl as a Stable Structural Mimic of Reactive Ferryl Intermediates in Mononuclear Nonheme-Iron Enzymes,” Inorg. Chem. 2017, 56, 13382-13389.

13. Mitchell, A. J.; Dunham, N. P.; Martinie, R. J.; Bergman, J. A.; Pollock, C. J.; Hu, K.; Allen, B. D.; Chang, W.-c.; Silakov, A.; Bollinger, J. M., Jr.; Krebs, C.; Boal, A. K. “Visualizing the Reaction Cycle in an Iron(II)- and 2-(Oxo)-glutarate-Dependent Hydroxylase,” J. Am. Chem. Soc. 2017, 139, 13830-13836.

Abstract:

Exposure to particulate matter pollution (PM, tiny particles suspended in air) ranks among the top ten global health risks. PM-induced oxidative stress has been suggested as a possible mechanism leading to their health effects. We conducted cellular and acellular measurements of PM-induced oxidative stress through systematic laboratory experiments and ambient field studies. Murine alveolar macrophages were used to measure intracellular reactive oxygen and nitrogen species (ROS/RNS) production, while dithiothreitol (DTT) was used to measure the chemical oxidative potential for each sample. Ambient samples (n = 104) were collected during multiple seasons from rural and urban sites around the greater Atlanta area as part of the Southeastern Center for Air Pollution and Epidemiology (SCAPE). For laboratory studies, SOA were generated from photooxidation of six commonly emitted volatile organic precursors, including isoprene, α-pinene, β-caryophyllene, pentadecane, m-xylene, and naphthalene. Laboratory experiments were conducted in the Georgia Tech Environmental (GTEC) facility under different conditions. For both ambient and laboratory samples, we found that cellular ROS/RNS production was highly dose-dependent, non-linear, and could not be represented by a single concentration measurement. For chemical oxidative potential, precursor identity influenced toxicity significantly, with isoprene and naphthalene SOA having the lowest and highest response, respectively. Both precursor identity and formation conditions influenced inflammatory responses. Several response patterns were identified for SOA precursors whose photooxidation products share similar carbon chain length and functionalities. A significant correlation between ROS/RNS levels and aerosol carbon oxidation state was also observed, which may have significant implications as ambient aerosols have an atmospheric lifetime of a week, over which oxidation state increases due to photochemical aging, potentially resulting in more toxic aerosols.

The human brain with about 10¹¹ neurons and about 10¹⁵ connections in addition to neuroglia which consists of even more numerous cells of various types is one of the most complex and fascinating systems in the universe. The connections between neurons are established through axons, which are long and often thin structures that carry electrical signals also called action potentials. Physiologic function relies on correct timing of the arrival of the electric signals and hence speed at which they travel along the axons, which, in turn, depends strongly on the diameter of the axon, which therefore must be precisely matched to its physiologic function.

The principal determinant of axon diameter in vertebrates are space-filling cytoskeletal polymers called neurofilaments (NFs). Morphometric studies have indeed established a direct correlation between NFs and axonal diameter. In addition to their space-filling role, NFs are also cargo of slow axonal transport and are in relentless but slow movement toward the nerve terminals. The focus of our collaborative research with the Brown-lab at Ohio State University is a new paradigm for the understanding of axon morphology that is rooted in the dual motile and architectural function of NFs as cargo of slow transport and space-filling structures. According to this view, axon caliber is emergent and dynamically determined by changes in the flow of NFs.  We combine fluorescent life imaging methods to characterize the dynamics of the cytoskeleton of the axon, with mathematical and computational modeling to understand how axon caliber is regulated, how morphological structures, such as constrictions at nodes of Ranvier or neurodegenerative disease related swellings, are formed.